Establishing a Mouse Model of Thin Endometrium

Summary

This article describes a detailed protocol for producing a reliable and reproducible thin endometrium with a very low mortality rate and minimal intrauterine adhesions by injecting 95% ethanol into the mouse uterus within 1-3 min.

Abstract

Thin endometrium (TE) has been widely recognized as a critical cause of infertility. However, the pathogenesis of TE remains unclear, and satisfactory treatment options are still urgently needed. Several animal models of TE have been developed, but the mouse model involving abdominal surgery and injection of 95% ethanol presents a formidable challenge due to the high mortality rate and risk of intrauterine adhesions if not performed correctly. Here, we describe a detailed protocol that produces reliable and reproducible TE with a very low mortality rate and minimal intrauterine adhesions by injecting 95% ethanol into the mouse uterus with varying infusion times. The results showed that all of the mice successfully developed TE with infusion times ranging from 1-3 min, characterized by a typical reduction in endometrial thickness and the number of glands, as well as excessive endometrial fibrosis. These findings suggest that this mouse model is suitable for studying thin endometrium and can serve as a platform for developing future TE treatments.

Introduction

Thin endometrium (TE) is a serious condition in obstetrics and gynecology that often affects women of childbearing age. TE is diagnosed when the endometrium thickness measures less than 7 mm on an ultrasound scan, accompanied by a normal uterine cavity, and is closely associated with pregnancy failure^1,2. It is estimated that approximately 1.5%-9.1% of women undergoing in vitro fertilization (IVF) treatment will experience TE, making it a growing challenge in reproductive medicine^3,4,5. The most common causes of TE include improper endometrium repair following surgical separation of intrauterine adhesions (IUA) and curettage, which are often accompanied by disrupted blood vessel distribution and sparse glands^6,7,8. To date, the cellular and molecular mechanisms underlying TE remain unclear. Recovery of the endometrium in TE patients is time-consuming, although estrogen treatment and low-dose aspirin therapy have been explored as potential interventions⁹. Therefore, an in-depth study of the pathogenesis of TE is the most direct approach to addressing the challenges of this condition. However, studies on the pathogenesis of TE rely on animal models, making the selection of an appropriate model crucial.

Most histopathological studies of thin endometrium (TE) have shown that impaired proliferation of epithelial cells and macrophages, decreased expression of ovarian steroid hormone receptors, excessive deposition of extracellular matrix, and cellular senescence are the most significant pathological features of TE^6,9,10. Currently, various rat models have been developed to mimic TE, including models induced by scratching^11,12,13, ischemia¹⁴, thermal injury¹⁵, and chemical injury^16,17,18,19. A rat model is induced by scratching the endometrium with a needle or catheter, which often results in intrauterine adhesions (IUA) rather than TE^{11,12,20,21,22}. An ischemia-induced TE model in rats has also been reported, where endometrial ischemia is achieved by performing bilateral uterine artery ligation, leading to reduced endometrial thickness. However, this method is time-consuming, requiring three months, and is not widely used for research¹⁴. In the thermal injury-induced model, an artificial insemination tube is used to infuse 85 °C preheated water into one side of the uterine horn through the confluence of the two sides, making it more complex than other methods¹⁵. The chemical-induced model involves injecting 95% ethanol into the exposed uterine horn to damage the entire endometrium. This model offers the advantages of low cost and a short experimental period for studying the pathological mechanisms and treatments in TE, but it is primarily used in rats rather than mice^16,17,23. Although various animal models exist, especially rat models, each has its limitations. They can only simulate certain aspects of TE, and few mouse models closely replicate the disease characteristics of TE while also being convenient and versatile for research.

In this context, we developed a novel thin endometrium (TE) mouse model by infusing 95% ethanol into the uterine cavity using a time-gradient approach with a modified method²⁴ (Figure 1). The results showed that all the mice successfully developed TE when the infusion time ranged from 1-3 min, displaying typical characteristics such as reduced endometrial thickness, gland reduction, and increased endometrial fibrosis. These findings suggest that our mouse model is a suitable tool for studying TE and can serve as a platform for developing future TE treatments.

Protocol

This study was approved by the Institutional Animal Care and Use Committee of Shenzhen Zhongshan Obstetrics & Gynecology Hospital (formerly Shenzhen Zhongshan Urology Hospital), where all animal experiments were conducted. Female C57BL/6 mice (age 6-8 weeks, weight 18-20 g) were used in this study. All animals were housed in a specific pathogen-free (SPF) environment in the same room and acclimatized for one week before the experiments in a room without specific pathogens at (22 ± 1) °C under a 12-h light/dark cycle. They were provided with free access to food and water. The details of the reagents and equipment used in this study are listed in the Table of Materials.

1. Identification of estrus

NOTE: The estrus phase in mice is comparable to the late luteal phase in humans, with a relatively thick endometrial lining at this time. Selecting mice in estrus for model induction ensures that they are in a relatively consistent physiological state. Additionally, the results of endometrial thinning are more pronounced when the endometrial thickness is relatively thick. For details on this procedure, refer to previously published reports^25,26.

1. Prepare 20 µL of saline using a pipette. Gently insert the tip of the pipette into the vaginal orifice to a depth of approximately 2 mm. Infuse 20 µL of saline into the vagina and flush it gently for 5 cycles.
2. Withdraw the saline and transfer it evenly onto a clean slide. Allow it to evaporate and dry naturally at room temperature.
3. Apply 30 µL of ethanol to each slide to fix the cells. Allow the ethanol to evaporate for 5 min until the slide is dry.
4. Apply 50 µL of crystal violet staining solution to each slide and stain the cells at room temperature for 10 min.
5. Rinse each slide with deionized water to remove excess dye and reduce non-specific staining.
6. Observe the cell morphology and determine the proportion of cell types under a microscope.
7. Observe the estrous cycles of mice twice consecutively to ensure stability. Select mice in the third estrus for the experiment.
   NOTE: The estrous cycle in mice typically lasts 4-5 days and is divided into four stages: proestrus, estrus, metestrus, and diestrus. Proestrus lasts about 24 h and is characterized by round to oval nucleated epithelial cells (Figure 2A). Estrus lasts between 12 h and 48 h and is characterized by predominately anucleated keratinized epithelial cells (Figure 2B). Metestrus lasts up to 24 h and is characterized by a mix of anucleated keratinized epithelial cells and neutrophils (Figure 3C). Diestrus lasts between 48 h and 72 h and is characterized by a combination of neutrophils, nucleated epithelial cells, and a few anucleated keratinized cells (Figure 2D).

2. Group designing

NOTE: To assess the impact of various durations of 95% ethanol infusion on the stability of the TE model, six treatment groups were established. One group underwent sham surgery without ethanol administration to serve as a control for the procedural effects. Mice with ethanol-induced TE were randomly divided into five treatment groups based on the infusion time (n = 5 per group) as follows:

1. 1-min group: Inject ethanol into the uterine cavity for 1 min.
2. 2- min group: Inject ethanol into the uterine cavity for 2 min.
3. 3- min group: Inject ethanol into the uterine cavity for 3 min.
4. 4- min group: Inject ethanol into the uterine cavity for 4 min.
5. 5- min group: Inject ethanol into the uterine cavity for 5 min.

3. Induction of TE model in mice

1. Anesthetize the mice using an anesthesia machine with 5% isoflurane and 100% oxygen for 3 min (following an institutionally approved protocol). Maintain anesthesia at 1%-3% isoflurane using a nose cone. Apply veterinary ointment to the mice's eyes to prevent dryness during anesthesia. Monitor the respiratory rate with a toe pinch throughout the procedure.
2. Use medical tape to secure the mice's limbs and keep them in a supine position on the operating table preheated with an electronic blanket. Gently pull the mice's tongue slightly to prevent airway obstruction.
   NOTE: Keeping mice warm during surgery helps maintain immune function and blood circulation, and prevents clotting. Additionally, pulling out the mice's tongues ensures clear respiratory tracts to avoid suffocation during anesthesia.
3. Apply animal hair removal cream to the inferior abdomen of the mice for 3 min. Remove the hair with clean wipes. Disinfect the area with 75% ethanol and iodophor.
4. Make a 1 cm incision in the abdomen, 1 cm from the urethral orifice. Use sterile scissors and forceps to extend the incision down to the peritoneal cavity.
5. Gently move the entrails aside with sterile forceps to locate the uterus and expose the right uterine horn. Apply hemostatic clamps at the proximal end near the cervix and the distal end near the ovary.
   NOTE: Clamping both ends of the uterine horn ensures no leakage during ethanol injection, preventing damage to the cervix, vagina, and ovary.
6. Instill approximately 50 µL of 95% ethanol²⁷ into the uterine cavity using a 25 G syringe, with the needle parallel to the direction of the uterine horn. Insert the needle into the uterine cavity at a 30-degree angle. Maintain the syringe for 1 min, 2 min, 3 min, 4 min, or 5 min, as appropriate. Withdraw the ethanol and flush the uterine cavity with sterile saline 5 times to remove any remaining ethanol. Similarly, instill an equal volume of sterile saline into the left uterine horn as a control, following the same procedures.
   NOTE: Ensure the syringe needle is parallel to the direction of the uterine horn. Avoid inserting the needle at an angle that is too steep to prevent it from traversing the entire uterine horn. Withdraw the ethanol as completely as possible to avoid persistent damage and experimental instability.
7. Remove the clamps and return the uterus and entrails to their original position.
8. Suture the peritoneum with 5-0 absorbable surgical sutures, followed by suturing the epidermis with 5-0 non-absorbable surgical sutures.
9. Disinfect the incision with 75% ethanol and iodophor, and then return the animals to their cage (Figure 2A-H).
10. Observe the mice at 9:00 and 15:00 each day and disinfect the incision with iodophor to ensure they are in good condition.

4. Sample collection

1. Euthanize the mice by cervical spine dislocation 7 days after 95% ethanol treatment (following an institutionally approved protocol).
2. Cut the sutures to expose the uterus using surgical scissors.
3. Carefully and slowly extract the entire uterus using surgical forceps. Detach the uterus from the surrounding entrails.
4. Immediately remove both uterine horns and divide each horn into 3-4 small segments.
5. Add 1 mL of 4% paraformaldehyde to a 1.5 mL microcentrifuge tube using a pipette. Preserve the uterine tissue in 4% paraformaldehyde for 24 h for fixation.

5. Tissue dehydration

NOTE: Tissue dehydration is achieved using an automatic tissue processor (see Table of Materials).

1. Place the formalin-fixed samples into tissue cassettes.
2. Transfer the tissue cassettes through a series of increasing ethanol concentrations to dehydrate the tissue as follows: 70% ethanol for 1 h, 85% ethanol for 1 h, 95% ethanol twice (1 h each), and 100% ethanol twice (1 h each).
3. Replace the ethanol with 100% xylene (135 min for the first immersion, 20 min for each subsequent immersion twice) to ensure that the paraffin penetrates the tissue.
4. Immerse the tissue cassettes in paraffin in an incubator at 60-65 °C (3 times, 140 min each), and keep the cassettes warm until embedding.

6. Paraffin embedding

NOTE: Perform these steps using a modular tissue embedding center (see Table of Materials).

1. Fill a wax mold with molten paraffin wax heated to 55 °C.
2. Use heated forceps to quickly transfer the infiltrated segments from the embedding cassette into the wax mold, ensuring that the cross-section of each segment rests parallel to the base of the mold.
3. Once the segments are properly positioned, place the bottom of the embedding cassette on top of the histology mold.
4. Place the mold directly on a cooling plate set to 4 °C for at least 20 min to allow the paraffin to solidify.
5. After the wax has solidified, remove the sample from the mold and use a paraffin trimmer to trim any excess wax around it.

7. Paraffin sectioning

1. Prepare a water bath at 42 °C. Place the paraffin blocks on a cold plate before cutting to allow them to cool and become easier to trim.
2. Clamp the paraffin-embedded specimen onto a rotary microtome with the tissue surface facing up.
3. Section 20 µm at a time to remove excess paraffin covering the tissue until the desired tissue plane is reached.
4. Cut 4 µm sections at a time.
5. Transfer the paraffin-sectioned ribbon to the 42 °C warm water bath for 2 min.
6. Pick up the sections from the water surface and place them onto a microscope slide after unfolding.
7. Dry the slides in a 60 °C incubator for 30 min.

8. Hematoxylin and eosin staining

NOTE: Hematoxylin and Eosin staining is performed using an automated slide stainer machine (see Table of Materials).

1. Prepare all reagents in advance before starting the staining procedure.
2. Soak the slides in 100% xylene for 10 min, 5 min, and 3 min, respectively.
3. Rehydrate the slides in a decreasing ethanol series in deionized water as follows: 100% ethanol, 95% ethanol (twice), and 80% ethanol, for 2 min each.
4. Rinse the slides in deionized water.
5. Stain the slides in hematoxylin solution for 10 min. Rinse the slides in deionized water for 1 min or until the water is no longer purple.
6. Dip the slides in 0.5% acid ethanol differentiation solution twice for 3-5 s each time. Rinse the slides in deionized water for 1 min.
7. Place the slides in a bluing buffer (1% ammonia solution) for 1 min. Rinse the slides in deionized water for 1 min.
8. Dehydrate the slides with 80% ethanol and then dip them in an eosin-Y working solution for 10 s.
9. Transfer the slides to 95% ethanol and rinse twice for 10 s each. Then, transfer the slides to absolute ethanol and dehydrate them twice for 10 s each.
10. Dip the slides in 100% xylene to clear the tissue twice, for 2 min each.
11. Place a coverslip over the tissue and seal it with natural resin.

9. Masson staining

1. Deparaffinize the tissue by soaking the slides in 100% xylene for 10 min, 5 min, and 3 min, respectively.
2. After deparaffinizing, soak the slides in a decreasing ethanol series in deionized water: 100% ethanol, 95% ethanol (twice), and 80% ethanol, for 2 min each.
3. Rinse the slides in deionized water.
4. Stain the slides in Weigert's iron hematoxylin solution for 5 min, then wash the slides in deionized water for 30 s.
5. Differentiate the colors of the stained tissue structures by incubating the slides in 1% hydrochloric acid alcohol for 15 s.
6. Rinse the slides in deionized water for 30 s.
7. Incubate the tissue in a Bluing Solution for 3-5 min.
8. Rinse the slides in deionized water for 30 s.
9. Place the slides in Ponceau-Acid Fuchsin Solution for 5-10 min.
   NOTE: Mix deionized water and weak acid solution in a ratio of 2:1 to prepare the weak acid working solution.
10. Rinse the slides with the weak acid working solution for 30 s.
11. Differentiate the tissue in the phosphomolybdic acid solution for 1-2 min, and then rinse the sections with a weak acid working solution for 30 s.
12. Stain the tissue sections with aniline blue solution for 1-2 min.
13. Dehydrate the tissue sections quickly in 95% ethanol for 2-3 s, followed by 100% ethanol twice for 5-10 s each. Clear the slides in xylene twice, for 1-2 min each.
14. Place a coverslip over the tissue and seal it with natural resin.
15. Drain excess resin and allow the slides to dry.

10. Imaging and analysis

1. Capture images of the sections using a slide scanner system.
   NOTE: Use a 4x or 10x objective for an overview of the sections; for more detailed images, use 20x to 100x objectives.
2. Perform quantitative analysis of endometrial thickness, the number of glands, and the extent of tissue fibrosis using the HALO image analysis platform.
   NOTE: Average thickness is obtained by measuring the vertical distance from the uterine cavity to the myometrium in 5 randomly selected fields of view.

11. Statistical analyses

1. Conduct statistical analyses using SPSS version 23.0 (SPSS Inc., Chicago, IL, USA).
2. Assess the statistical significance of experimental differences between the control and model groups by first performing a normal distribution test.
3. Present data that are normally distributed as mean ± SEM.
4. Analyze data from different treatment groups using one-way ANOVA. Adjust for multiple comparisons of the number of tested alleles in each locus using the Bonferroni method. Consider a P-value <0.05 as statistically significant.

Results

The key features of thin endometrium (TE) are decreased endometrial thickness and glandular density, along with increased endometrial fibrosis. This method successfully replicated these characteristics in the model mice. Data analysis revealed a significant decrease in endometrial thickness in the 1-min group (222.3 µm ± 13.96 µm vs. 359.2 µm ± 12.41 µm, P < 0.05), the 2-min group (168.7 µm ± 17.57 µm vs. 359.2 µm ± 12.41 µm, P...

Discussion

TE is characterized by insufficient cell proliferation and dysfunctional cells, closely linked to infertility, recurrent miscarriage, and placental abnormalities^2,3. Unfortunately, there is currently no effective therapy for TE. Animal models play a crucial role in studying this condition. Between 2014 and 2024, rats were used as model organisms in 16.4% of 208,000 studies (34,200 studies) and mice in 22.7% (47,300 studies). The increasing use of mice in experime...

Materials

Name	Company	Catalog Number	Comments
Anesthesia Machine	RWD Life Science	R530	Mobile inhalation anesthesia machine for small animals
0.9% saline	Hubei Kelun Pharmaceutical	C230817A2	10 mL, medical injection
75% ethanol	LIRCON	6303060031	500 mL, disinfectant reagent
95% ethanol	Guangzhou Chemical Reagent Factory	64-17-5	500 mL, chemical reagent
Absorbable sutures	Jinhuan Medical	CR537	Thickness: 5-0; Length: 90 cm
Aqueous ammonia	MilliporeSigma	1336-21-6	1000 mL, chemical reagent
Automatic Tissue Processor	Leica	TP1020	100 embedding boxes can be processed at one time
C57BL/6 mice	Experimental Animal Center of Southern Medical University
Eosin-Y	BASO	BA4024	1000 mL, used for the staining of paraffin sections, frozen sections, etc
Haematoxylin	BASO	BA4041	1000 mL, used for the staining of paraffin sections, frozen sections, etc
HALO Image Analysis Platform	Indica labs		The instrument features ease-of-use and scalability, powerful analytical capabilities, and the fastest processing speed
Hemostatic clamps	HUAYON	18-5021	1.8 cm in total length with 0.7 cm jaw
Hemostatic forceps	HUAYON	18-5020	10 cm in total length
HistoCore Rotary Microtome	Leica	149BIO000C1	Slice thickness ranges from 1 to 60 μm
Indorphor	ADF	1005	500 mL, disinfectant reagent
Isoflurane	RWD Life Science	R510-22-10	100 mL, active ingredient 100% isoflurane
Masson's trichrome staining	SOLARBIO	G1340	7 × 100 mL for 100 tests
Microscope slide	Gene Tech	GT100511	Length: 75 cm; Width: 25 cm
Modular Tissue Embedding Center	Leica	EG1150 C	The instrument contains a cold stage and a heated paraffin distribution module, providing flexibility for the embedding work
Natural resin	SAKURA	4770	Resin-coated film, Suitable for histology staining
Olympus SLIDEVIEW VS200	PANOVUE	VS200	The instrument captures high-quality virtual slide images and enables advanced quantitative image analysis
Paraformaldehyde fix solution	Servicebio	G1011	500 mL, universal tissue fixative (neutral)
Surgical forceps	HUAYON	18-1300	2.2 mm straight, 12.5 cm wide
Surgical scissors	HUAYON	18-0110	10 cm, stainless steel surgical scissors
Syringe	Kindly	60017031	1 mL, disposable sterile syringe with needle
Tissue cassettes	CITOTEST	80106-1100-16	White; flow-through slots; 0ne-piece integral lid; labeling areas are located on three sides
Tissue-Tek Prisma Plus	SAKURA	DRS-Prisma-P-JCS	The processing capacity is 60 slides at one time
Xylene	Guangdong Guanghua Sci-Tech	1330-20-7	1000 mL, organic solvent